Lyophilization induces physicochemical alterations in cryptococcal exopolysaccharide

Carbohydrate Polymers 291 (2022) 119547

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Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol

Lyophilization induces physicochemical alterations in
cryptococcal exopolysaccharide
Maggie P. Wear a, *, Audra A. Hargett b, John E. Kelly c, Scott A. McConnell a,
´n I. Freedberg b, Ruth E. Stark c, d, Arturo Casadevall a
Conor J. Crawford a, 1, Daro
a

W. Harry Feinstone Department of Molecular Microbiology and Immunology, Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA
Laboratory of Bacterial PSs, Office of Vaccines Research and Review, Center for Biologics Evaluation and Research, U.S. Food and Drug Administration, Silver Spring,
MD, USA
c
Department of Chemistry and Biochemistry, The City College of New York and CUNY Institute for Macromolecular Assemblies, New York, NY, USA
d
Ph.D. Programs in Chemistry and Biochemistry, The City University of New York, New York, NY, USA
b

A R T I C L E I N F O

A B S T R A C T

Keywords:
Cryptococcus
NMR
DLS

TEM
Exopolysaccharide
Lyophilization

Microbial polysaccharide characterization requires purification that often involves detergent precipitation and
lyophilization. Here we examined physicochemical changes following lyophilization of Cryptococcus neoformans
exopolysaccharide (EPS). Solution 1H Nuclear Magnetic Resonance (NMR) reveals significant anomeric signal
attenuation following lyophilization of native EPS while 1H solid-state Nuclear Magnetic Resonance (ssNMR)
shows few changes, suggesting diminished molecular motion and consequent broadening of 1H NMR poly­
saccharide resonances. 13C ssNMR, dynamic light scattering, and transmission electron microscopy show that,
while native EPS has rigid molecular characteristics and contains small, loosely packed polysaccharide assem­
blies, lyophilized and resuspended EPS is disordered and contains larger dense aggregates, suggesting that
structural water molecules in the interior of the polysaccharide assemblies are removed during extensive
lyophilization. Importantly, mAbs to C. neoformans polysaccharide bind native EPS more strongly than lyophi­
lized EPS. Together, these observations argue for caution when interpreting the biological and immunological
attributes of polysaccharides that have been lyophilized to dryness.

1. Introduction
Cryptococcus neoformans is protected from the environment and in
mammalian infection by a complex polysaccharide capsule. This capsule
is a highly hydrated structure and as such, it has a refractive index that is
very similar to water, making it difficult to visualize. In the environ­
ment, the capsule protects the fungal cell from amoeba predation and
dehydration (Aksenov, Babyeva, & Golubev, 1973; Steenbergen, Shu­
man, & Casadevall, 2001). During mammalian infection, the capsule
protects the fungal cell from phagocytic cells (Kozel, Pfrommer, Guer­
lain, Highison, & Highison, 1988). Additionally, during cryptococcal
infection large quantities of cryptococcal polysaccharide are shed into
tissue, and this material interferes with effective immune responses
(Kang et al., 2004; A Vecchiarelli, 2000), overall exacerbating the

infection. Detection of cryptococcal polysaccharide in blood and cere­
brospinal fluid also provides physicians with important diagnostic and

prognostic information for C. neoformans disease.
The last five decades have witnessed significant efforts to understand
the cryptococcal capsular architecture and yielded important biophysi­
cal, chemical, and structural information about the polysaccharide
capsule. The dominant polysaccharide component of the C. neoformans
capsule is glucuronoxylomannan (GXM). Cryptococcal EPS structure has
been inferred from light scattering analysis of shed exopolysaccharide
(EPS), revealing GXM to be large dense branched polymers (Cordero,
Frases, Guimară
aes, Rivera, & Casadevall, 2011) that self-aggregate
(Nimrichter et al., 2007) to form rosette-like condensed structures
(Cordero et al., 2011; McFadden, De Jesus, & Casadevall, 2006; Nim­
richter et al., 2007) 1700–7000 megadaltons in size (McFadden et al.,
2006). The GXM polymer consists of an α-(1,3)-mannose backbone with
a β-(1,2)-glucuronic acid branch at every third mannose and varied
β-(1,2)- and β-(1,4)-xylose branches from the mannose backbone
(Bhattacharjee, Bennett, & Glaudemans, 1984; Cherniak & Sundstrom,

* Corresponding author.
E-mail address: (M.P. Wear).
1
Current address: Department of Biomolecular Systems, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany.
/>Received 16 March 2022; Received in revised form 14 April 2022; Accepted 25 April 2022
Available online 29 April 2022
0144-8617/© 2022 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY license ( />

M.P. Wear et al.

Carbohydrate Polymers 291 (2022) 119547

1994; Turner & Cherniak, 1991). The varied xylosylation results in tri­
mannose repeat motifs, seven of which have been described for GXM
(Cherniak, Valafar, Morris, & Valafar, 1998). In previous studies, the
cryptococcal EPS has been isolated using purification steps that require
precipitation with cetyl trimethylammonium bromide (CTAB) detergent
followed by ethanol precipitation, ultrasonication, dialysis, lyophiliza­
tion, and base treatment to remove O-acetylation (Cherniak et al.,
1998). The arrangement of these trimannose motifs into higher order
polysaccharide structures has largely remained beyond the reach of
technologies for polymer purification and analysis. Changes to the
overall polysaccharide organization and structure, depending upon the
preparation technique, were evidenced by Circular Dichroism (CD)
peaks of higher molar ellipticity in the far-UV region (Frases, Nim­
richter, Viana, Nakouzi, & Casadevall, 2008). Additionally, ultrafiltra­
tion without lyophilization resulted in 14-fold less dense preparations
than CTAB precipitation and lyophilization, suggesting that ultrafiltered
EPS is organized differently from CTAB-precipitated and lyophilized
samples (Nimrichter et al., 2007). A recent publication postulated that
all natural polysaccharides may have physicochemical differences
depending upon the method of preparation and that these physico­
chemical differences translate into functional effects (Yi, Xu, Wang,
Huang, & Wang, 2020).
Here we present evidence of physicochemical alterations to crypto­
coccal EPS induced by lyophilization to the point of dryness, a technique
relied upon for non-sterile EPS isolation. Solution 1H NMR spectra of
native EPS contain peaks in the SRG region (5.0–5.4 ppm) are consistent
with GXM; whereas after the sample was lyophilized to dryness and

solvated with water, peaks in the SRG region were significantly atten­
uated or lost. However, magic-angle spinning solid-state 1H NMR (MAS
ssNMR) spectra indicate that the native EPS and lyophilized samples
contained similar material. Therefore, we hypothesize that the attenu­
ation of solution NMR signals after EPS lyophilization originates from a
change in physicochemical properties rather than changes to chemical
structure. In support of this hypothesis, contrasting physical measure­
ments showed that the lyophilized EPS differed from the parent native
material in several ways: lyophilized EPS is larger, more mobile, more
disordered, and was less reactive with mAbs to GXM. Together, our
findings implicate alterations after lyophilization of these polymers. As
the majority of published studies on C. neoformans EPS rely on lyophi­
lized material, it is essential to consider the impact of these observed
differences on the interpretation of previous structural and immuno­
logical studies and the design of future investigations that can deepen
our understanding of the role of these PS structures in fungal infection.

were collected at 60 ◦ C, with 64 scans and a free induction decay size of
84336 points. Standard Bruker pulse sequences were used to collect the
1D data (p3919gp and zggpw5). Data were processed in Topspin (Bruker
version 3.5) by truncating the FID to 8192 points using a squared cosine
bell window function and zero filling to 65536 points.
Lyophilized samples were dissolved in deuterated water to a con­
centration of 50 mg/ml or greater. Native samples were diluted by
adding 300 μl of D2O to 200 μl of sample. All NMR samples contained
DSS-d6 for chemical shift calibration and peak intensity comparisons.
2.3. Dynamic light scattering
Measurement of EPS particles by DLS was performed with a Zeta
Potential Analyzer instrument (Brookhaven Instruments). The particle
sizes in the suspension were measured for native samples as well as

lyophilized and rehydrated samples at different time points during a
period of 28 days. Data are expressed as the average of 10 runs of 1-min
data collection each. The multimodal size distributions of the particles
were obtained by a non-negatively constrained least squares algorithm
based on the intensity of light scattered by each particle. The multi­
modal size distributions of particles from each sample were graphed for
comparison.
2.4. ELISA
For capture ELISA, Microtiter polystyrene plates were coated with
goat anti-mouse IgM at 1 μg/ml (SouthernBiotech, Birmingham, AL) and
then blocked with 1% BSA blocking solution. 2D10, a murine anti-GXM
IgM, was subsequently added at 10 μg/ml as the capture antibody. Next,
lyophilized or native wEPS samples were added to each half of the plate
and serially diluted. 18B7, a murine anti-GXM IgG1, was added at 10 μg/
ml and serially diluted in the opposite direction as the GXM dilution. The
direct ELISA was performed by coating plates directly with antigen
(native or lyophilized EPS) at 1 μg/ml, followed by 18B7 at 5 μg/ml to
each well. For both, the assays were developed by sequential addition of
goat anti-mouse IgG1 conjugated to alkaline phosphatase at 1 μg/ml and
1 mg/ml p-nitrophenol phosphate substrate. The absorbance of each
well was measured at 405 nm after a short incubation at 37 ◦ C. Between
each step of the ELISAs, the plate was incubated for 1 h at 37 ◦ C and
washed three times in 0.1% Tween 20 in Tris-buffered saline.
2.5. Solid-State NMR

2. Materials and methods

Partially dehydrated samples were prepared by lyophilizing a 5-mL
EPS solution for 18 h to obtain 309 mg of a ‘cookie dough’ material
that was packed into a 3.2-mm OD ssNMR rotor. Partially rehydrated

samples were prepared by adding 0.08 ml of water to 211 mg of fully
dried EPS powder, matching the weight percent of the ‘cookie dough’
and yielding 297 mg of a ‘sticky batter.’ NMR spectra were acquired
with a Varian (Agilent) DirectDrive2 spectrometer operating at a 1H
frequency of 600 MHz and using a 3.2-mm T3 HXY Magic Angle Spin­
ning (MAS) probe (Agilent Technologies, Santa Clara, CA). These data
were acquired on 34.1 and 39.7 mg, respectively, of concentrated
(partially dehydrated) and lyophilized (partially rehydrated) samples
using a spinning rate of 15.00 ± 0.02 kHz and a nominal temperature of
25 ◦ C. The 1H spectra were obtained with a single 90◦ pulse, whereas 13C
spectra used either 1-ms 1H–13C cross polarization (CP) with a 10%
ramp of the 1H power and 3 s between data acquisition sequences or
direct polarization (DP) with a 2-s recycle delay. 1H decoupling with a
radiofrequency field of 109 kHz was applied during signal acquisition
with the small phase incremental alternation method (Fung, Khitrin, &
Ermolaev, 2000) . After apodization of the data with a decaying expo­
nential function to improve the signal-to-noise ratio and Fourier trans­
formation, the spectra were referenced to H2O at 4.8 ppm.

2.1. Fungal growth and exopolysaccharide isolation
C. neoformans serotype A strain H99 (ATCC 208821) cells were
inoculated in Sabouraud rich medium from a frozen stock and grown for
two days at 30 ◦ C with agitation (150 rpm). Capsule growth was induced
by growth in chemically defined media (7.5 mM glucose, 10 mM
MgSO4, 29.4 mM KH2PO4, 6.5 mM glycine, and 3 μM thiamine-HCl, pH
5.5) for 3 days at 30 ◦ C, with agitation (150 rpm). The supernatant was
isolated from cells by centrifugation (4,000 xg, 15 min, 4 ◦ C) and sub­
sequently sterilized by passing through a 0.45 μm filter. Native samples
were concentrated while lyophilized samples were freeze dried to
complete dryness, defined by no change in mass with time, for an

average of 5 to 7 days.
2.2. Solution NMR
1D 1H NMR data were collected on either of two spectrometers: a
Bruker Avance II (600 MHz), equipped with a triple resonance, TCI
cryogenic probe and Z-axis pulsed field gradients or a Bruker Avance III
HD (700 MHz), equipped with an XYZ gradient TCI cryoprobe. Spectra
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Carbohydrate Polymers 291 (2022) 119547

2.6. Negative staining with uranyl acetate and transmission electron
microscopy

dough and (b) resuspending dry lyophilized EPS with the quantity of
water to match that remaining in (a). 1D 1H MAS ssNMR showed that
both the native (concentrated, partially dehydrated) and lyophilized
(partially rehydrated) H99 wEPS samples display the same set of reso­
nances (Fig. 1E, F), though the relative peak intensity for the 6.8-ppm
signal is notably altered and the SRG signals are overlapped by the
solvent in both preparations. These observations suggest that most of the
wEPS material present in the resuspended samples maintained its
chemical structure but was insufficiently solvated to allow the molecular
moieties to become more mobile and thereby more easily observable in
the solution-state NMR spectra. In both the native (concentrated) and
lyophilized (partially rehydrated) samples, wEPS was solvated suffi­
ciently to be observed by NMR when MAS was used to average out many
of the anisotropic spin interactions described above. To our knowledge,

these are the first NMR findings that explore the impact of dehydrationrehydration procedures on cryptococcal polysaccharide structure.

Samples (10) were adsorbed to glow discharged (EMS GloQube)
ultra-thin (UL) carbon coated 400 mesh copper grids (EMS CF400-CuUL), by floatation for 2 min. Grids were quickly blotted then rinsed in
3 drops (1 min each) of TBS. Grids were negatively stained in 2
consecutive drops of 1% uranyl acetate with tylose (UAT), then quickly
aspirated to get a thin layer of stain covering the sample. Grids were
imaged on a Hitachi 7600 TEM (or Philips CM120) operating at 80 kV
with an AMT XR80 CCD (8 megapixel).
3. Results
3.1. NMR signals are attenuated or absent in rehydrated
exopolysaccharide samples

3.3. Solid-state 13C NMR reveals differences in molecular mobility of
polysaccharide in the wEPS samples

Samples of C. neoformans (H99) whole exopolysaccharide (wEPS)
were processed only by sterile filtration (0.22 μm). We refer to this
sample as native. Half of this native sample was then lyophilized (~5 d)
until the dry weight did not change and solvated with water. We
weighed the sample before (97.80 g, average n = 3) and after (0.82 g,
average n = 3) lyophilization. The loss of mass as a result of lyophili­
zation is (average n = 3, 96.79 g) 99.16% of the total mass. This result is
consistent with a previous study using γ-irradiation to strip the outer
capsule, which reduced the cell pellet volume by 85%, suggesting the
majority of capsular polysaccharide mass is water (Maxson, Cook,
Casadevall, & Zaragoza, 2007).
Following this analysis, both samples were examined by 1D 1H NMR
in solution. The solution 1H NMR spectrum of the native sample showed
a peak set in the structural reporter group (SRG) region (5.0–5.4 ppm),

as defined by Cherniak and colleagues (Cherniak et al., 1998) (Fig. 1A).
However, when we examined the same material that had been lyophi­
lized and solvated with water, we found that not all material went into
solution. Additionally, the peaks in the SRG region were significantly
diminished in intensity or were lost (Fig. 1B) (Cherniak et al., 1998),
even after attempting to re-solubilize the EPS at 37 ◦ C for 14 days with
agitation (Fig. 1C). While not quantitative due to a lack of baseline peak
resolution, overlays of the three spectra, normalized by setting the DSS
signal to 1.0, demonstrated that peaks in the SRG region at 5.35, 5.22,
and 5.18, were reduced by approximately 60, 80, and 30% respectively,
in the lyophilized sample (Fig. 1D). Interestingly, two distinct biological
isolates of H99 EPS treated the same way contained peaks at similar
resonance frequencies in the SRG, but the intensity of these peaks
differed, partially due to variation in the amount of O-acetylation
(Fig. S1). However, both sample sets show decrease in peak signal in­
tensity after lyophilization. While there seems to be a greater level of
diversity in the polysaccharide of H99 than observed for other strains,
the reduction in signal was consistent between samples. C. neoformans
EPS is generally understood to be solvated with water but we wondered
if more hydrophobic solvents could reconstitute the lyophilized material
more effectively. We attempted to recover the missing NMR resonances
by dissolution of lyophilized wEPS in acetonitrile and dimethyl sulfoxide
but neither was superior to water at restoring the signals of the SRG
region.

A confirmation of the physicochemical rationale for the 1H NMR
observations and a more detailed comparison of the native (concen­
trated) and lyophilized (partially rehydrated) EPS materials were
available from a follow-up set of 13C ssNMR experiments. To probe the
impact of hydration at particular molecular sites of the EPS polymers,

we acquired both cross polarization magic angle spinning (CPMAS) 13C
ssNMR (to favor detection of rigid and protonated polymeric moieties)
and direct polarization magic angle spinning (DPMAS) 13C ssNMR with a
short (2-s) delay between successive spectral acquisitions (to ensure
inclusion of mobile and disordered chemical groupings in the spectra).
Whereas the CPMAS spectra display no EPS signals for either partially
dehydrated or partially rehydrated samples (Fig. S2), the DPMAS spectra
(Fig. 2) reveal relatively sharp resonances from the mobile glycan
groups (~62–105 ppm) in both native (concentrated) and lyophilized
(partially rehydrated) samples, but no significant contributions from
alkene or carboxyl carbons with chemical shifts above 110 ppm.
Notably, the major glycan resonances between ~62 and 105 ppm are
sharper and thus better resolved in the lyophilized (partially rehydrated)
sample, indicating more complete solvation and motional averaging of
the polysaccharide structures. The mobility that yields resolved 1H and
13
C NMR spectra under magic-angle spinning acquisition conditions can
be attributed to the hydrophilic nature of the sugar ring structures.
3.4. Electron microscopy of EPS shows rosette-like assemblies in
rehydrated lyophilized sample
To further investigate the effects that lyophilization might have on
EPS, we turned to Transmission Electron Microscopy (TEM) to examine
the architecture of the EPS samples. This analysis shows that the native
EPS is less dense and contains vesicles (Fig. 3A). The presence of vesicles
is not surprising since these are shed by C. neoformans during capsule
growth (Oliveira et al., 2010; Rodrigues et al., 2008) and would be
retained by the filtration step. In contrast, no vesicles were observed in
the lyophilized and reconstituted samples, possibly reflecting collapse of
these structures during the drying procedure (Merivaara et al., 2021).
The lyophilized and reconstituted material does contain dense, rosettelike assemblies, similar to those observed previously for cryptococcal

capsular polysaccharide isolations and glycogen (Fig. 3B) (Childress,
Sacktor, Grossman, & Bueding, 1970; Cordero et al., 2011).

3.2. Solid-state 1H NMR displayed the same chemical reporter groups in
native or lyophilized EPS
We then turned to solid-state 1H NMR accompanied by magic-angle
spinning (MAS) for partially hydrated wEPS samples, endeavoring to
average the orientation-dependent chemical shift tensors to their liquidstate values and remove 1H–1H dipolar couplings between pairs of
nuclear spins that are situated within ~1 nm of one another (Laws,
Bitter, & Jerschow, 2002). The ssNMR samples were made by (a)
concentrating a native EPS solution to a consistency resembling cookie

3.5. Dynamic light scattering shows size differences as a function of
solubilization time
Dynamic light scattering (DLS) revealed that the average effective
diameter for particles in the native wEPS preparation were ~ 115 nm
but after lyophilization these increased in size (~300 nm and ~ 8500
3


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Carbohydrate Polymers 291 (2022) 119547

A. H99 EPS Native

5.4

5.3

5.2
[1H] ppm

8

B. H99 EPS Lyophilized - 0 time

5.1

6

5.0

[1H] ppm

5.4

4

5.3

8

2

5.2
[1H] ppm

5.1

6

D. SRG Region Overlay

C. H99 EPS Lyophilized - 14 days resuspended

5.0

[1H] ppm

4

2

H99 EPS Lyophilized - 14 day solubilized
H99 EPS Lyophilized - 0 time
I
H99 EPS Native

II

III

5.4

5.3

8

E.

5.2
[1H] ppm

5.1

6

5.0

[1H] ppm

4

2

5.4

H99 EPS Native (concentrated)

5.3

5.2
[1H] ppm

5.1

5.0

F. H99 EPS Lyophilized (partially rehydrated)

[1H] ppm

[1H] ppm

Fig. 1. Effects of lyophilization on NMR signals of C. neoformans EPS. One-dimensional 1H solution NMR spectra and insets expanded vertically by factors of 10 at 60
◦
C for a native (A) preparation compared with preparations which were lyophilized and solvated with water at time 0 (B) and after 14 days (C); the three spectra are
overlayed in (D). SRG region peaks which were integrated indicated as I, II, and III as the motif they belong to is unknown. Peak integrals for the SRG region of the
solution-state spectra were compared by setting the respective DSS signals to 1.0. One-dimensional 1H solid-state NMR (ssNMR) spectra obtained at room tem­
perature with 15-kHz magic-angle spinning are shown for native (E: concentrated, partially dehydrated) and lyophilized (F: partially rehydrated) samples,
normalized according to sample mass. The chemical shifts of the solution- and solid-state spectra were referenced to DSS at 0.0 ppm and water at 4.8 ppm,
respectively. The sharp peaks in the ssNMR at 2.9 and 4.8 ppm are attributed to glycine and water, respectively.

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Carbohydrate Polymers 291 (2022) 119547

Fig. 2. Effects of lyophilization on solid-state 13C NMR spectra of EPS. 150 MHz 13C NMR spectra of C. neoformans EPS samples obtained with 15 kHz magic-angle
spinning (MAS), comparing samples that were native (partially dehydrated) (left) and lyophilized (partially rehydrated) (right). DPMAS experiments with short (2 s)
delays between successive cycles of signal acquisition, favoring carbon moieties that tumble rapidly in many directions. Sharp resonances at 40 and 170 ppm are
attributed to glycine in the culture media. (No EPS signals are observed in CPMAS experiments that favor rigid carbon moieties with nearby hydrogens.)

4. Discussion

nm) and cover a wider size range (Fig. 3C), consistent with reported
sizes for EPS particles (Cordero et al., 2011; McFadden et al., 2006;

Nimrichter et al., 2007). Over the course of 28 days in solution (D2O),
the effective diameter decreased (~950 nm with smaller particles)
(Fig. 3C).

Historically, analysis of the cryptococcal capsule has relied on the
examination of the shed EPS polymers. These analyses indicate that EPS
is composed of large, dense, branched polymers (Cordero et al., 2011;
McFadden et al., 2006; Nimrichter et al., 2007). Techniques for the
isolation of EPS have evolved since these initial analyses, however, the
maintenance of sterile sample preparations is challenging, resulting in
the use of lyophilization for long term polysaccharide sample storage.
There are indications in the literature that cryptococcal polysaccharide
is altered by the method of isolation (Frases et al., 2008; Nimrichter
et al., 2007; Rodrigues, Fonseca, Frases, Casadevall, & Nimrichter,
2009). Light scattering measurements of EPS samples reveal nine-fold
larger particles when precipitation with the cationic detergent CTAB is
utilized compared to isolation by ultrafiltration show (Nimrichter et al.,
2007), (Frases et al., 2008). Some investigators have stated outright that
the structure of EPS varies by method of preparation as well as that
CTAB isolation alters the secondary structure of polysaccharide
(Rodrigues et al., 2009). It is of utmost importance to define the impact
of polysaccharide preparation on the physicochemical and antigenic
properties of EPS, as our understanding of the immunoregulatory role of
cryptococcal EPS is largely derived from analysis of purified poly­
saccharide on immune cells.
One of the functions of the capsule is to protect the fungal cells from
dehydration (Aksenov et al., 1973). GXM, the predominant poly­
saccharide of the cryptococcal capsule, derives its hydrophilic nature
from its components – mannose, xylose, and glucuronic acid – as well as
the water coordination necessary to maintain the divalent cation bridges

formed between glucuronic acid residues (Nimrichter et al., 2007). To
appreciate the necessity of water to both the capsule and its composite
polymers, we can examine the sheer quantity of water present. As noted
above, when samples are weighed prior to and after lyophilization,
water makes up a significant proportion of the mass (98% in this study
and 85% in the gamma irradiation study). The additional data presented
here, including EM, DLS, solution NMR and solid-state NMR imply that
there may be internal or structural water molecules that are necessary
for the overall structure and organization of the polysaccharide assem­
bly. This suggests that water is critically important to the threedimensional structure of cryptococcal polysaccharide, not only form­
ing a hydration shell, but including structural waters. Our observations
indicate that after lyophilization the polysaccharide can be partially
solvated (hydration shell) but does not allow for the incorporation of
these structural waters.

3.6. ELISA uncovers antigenic differences between native and lyophilized
EPS
To examine how native and lyophilized EPS are bound by mono­
clonal antibodies (mAbs) to GXM, we performed capture ELISA, a
standard assay for determining GXM concentration in a sample
(Mukherjee & Casadevall, 1995). Fig. 4A and B show that mAbs to GXM
binds more strongly to native than lyophilized wEPS. This finding is
consistent with previous observations comparing CTAB- to
ultrafiltration-prepared EPS, wherein ultrafiltered EPS samples bound
both 12A1 and 18B7 better than CTAB precipitated EPS in direct ELISAs
(Nimrichter et al., 2007). These results suggest that there are more
available epitopes in the native polysaccharide than in wEPS that has
been lyophilized and resuspended.
3.7. Proposed model of dehydration-rehydration effects
Previous work by Cordero et al. showed that both EPS and CPS that

were lyophilized and rehydrated had hydrodynamic properties consis­
tent with a rosette-like condensed conformation (Cordero et al., 2011).
Similar aggregates can be visualized on the surface of cryptococcal cells
after dehydration imaged by scanning electron microscopy (SEM) and as
secreted particles by transmission electron microscopy (TEM) (Cordero
et al., 2011). These condensed formations have a higher density at the
core and more dispersed radial polymers. CTAB-precipitated EPS is also
more dense (14-fold) than the ultrafiltered native material (Nimrichter
et al., 2007). When these observations are considered in light of the
observations presented here, the suggested condensed conformations for
GXM are consistent with our lyophilized and resuspended material, but
not with native materials (Fig. 5), though other forms cannot be ruled
out. Prior to lyophilization, GXM polymers shed into solution as EPS by
C. neoformans are small, rigid, ordered, and hydrated. Lyophilization
results in the adoption of mobile, disordered, dense, and aggregated
architectures. While more time in solution may eventually restore these
polymers to resemble the native polysaccharide more closely, the
monthlong incubation time used in our study was insufficient to return
them to their native state (Fig. 5).
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Carbohydrate Polymers 291 (2022) 119547

A. Native EPS

100 nm

100 nm

B. Lyophilized EPS

100 nm

100 nm

C. DLS Particle Size

(caption on next page)
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Carbohydrate Polymers 291 (2022) 119547

Fig. 3. C. neoformans EPS undergoes biophysical changes over time in solution. Native EPS (A) and lyophilized and reconstituted EPS (B) samples were examined by
transmission electron microscopy (TEM) with negative staining at two levels of magnification. Native EPS contains a few aggregates as well as extracellular vesicles
(EVs), indicated with red arrows, while lyophilized and resuspended EPS contains many dense rosette-like structures previously reported for C. neoformans poly­
saccharides and glycogen (Cordero et al., 2011). Native EPS and lyophilized and reconstituted samples kept in solution for 7, 14, or 28 days were examined by DLS
(C). Lyophilized and resuspended samples have a larger particle size than native samples, judged by autocorrelation intensity of the scattered light as a function of
particle diameter.

Fig. 4. Lyophilization of C. neoformans EPS alters biological functions. Native EPS and lyophilized and resuspended samples were assayed for anti-GXM mAb binding
by capture ELISA. Binding curves (A) of serially diluted mAb 18B7 as a function of EPS concentration in capture ELISA. Native EPS generally binds more strongly to
the 2D10/18B7 capture/detection mAb pair than lyophilized EPS at a given mAb/antigen concentration. Statistical analysis (B) of binding in a capture ELISA doublearray assay varying both EPS and mAb concentration shows that native EPS is statistically significantly better bound by anti-GXM mAbs than lyophilized EPS. **** pvalue < 0.0001.

In this work, we examine an observed difference between native and

lyophilized EPS samples in which we noted the attenuation of anomeric
carbon signals in 1H solution-state NMR after lyophilization. However,
ssNMR 1H spectra show no significant difference between the signals in
the native and lyophilized samples, suggesting that the signal attenua­
tion in solution was due not to a chemical change, but to incomplete
solvation and subsequent failure to restore the polysaccharide assembly
to its native state (El Hariri El Nokab & van der Wel, 2020). The phys­
iochemical alterations resulting from loss of water could include
increased molecular size, decreased molecular mobility, and limited
angular excursions, which in turn would enhance nuclear spin relaxa­
tion and broaden the resonances to the point that individual signals
would appear to vanish (Ghassemi et al., 2021). Further comparison of
the two samples by TEM and DLS indicates that lyophilized and resus­
pended samples are larger, more mobile, and disordered. While most of
these observations are expected, the increased mobility runs counter to
the condensed conformations observed in the lyophilized sample. We
would note that the 13C ssNMR of lyophilized (partially rehydrated)
samples exhibit greater flexibility than the native (concentrated) sam­
ples, but neither of these states exhibits the rapid isotropic motions that
would yield well-resolved solution-state NMR spectra (Kelly, Chrissian,
& Stark, 2020).
Previous reports suggest that inter-polymer interactions occur
through divalent cation bridges formed between glucuronic acid resi­
dues of independent polymer. One interpretation of this effect is that the
solvation of these divalent cation bridges proceeds slowly over time.
However, we did not observe lyophilized molecules returning to their
pre-lyophilization, native, state after a month in solution. This may be
due to incomplete hydration, loss of mannose O-acetylation, or incom­
plete solvation wherein some polymers are recalcitrant to reconstitu­
tion. It is possible that the application of other conditions such as higher

temperature and/or different solution conditions (pH or electrolyte
concentrations) could return the polymers to their native state.
Although these physicochemical alterations to cryptococcal EPS are
interesting in their own right, we also observed functional changes as

suggested by Yi et al (Yi et al., 2020). C. neoformans EPS has been shown
to mediate numerous deleterious effects on host immune function (Anna
Vecchiarelli et al., 2013), which presumably result from the interaction
of carbohydrates with cellular receptors. Alterations in antigenic prop­
erties can be inferred from differences in mAb reactivity observed by
capture ELISA. Our observed reduction in binding of the lyophilized
sample by capture ELISA suggests decreased epitope prevalence and/or
accessibility, revealing that the physicochemical alterations effected on
the polysaccharide by lyophilization have a functional impact. Antibody
interactions may require a specific polysaccharide arrangement that is
altered by lyophilization, suggesting the need to revisit these observa­
tions with native material. Interestingly, although mAbs 2D10 and
18B7, the capture and detection mAbs used in this experiment, respec­
tively, were raised against CTAB-prepared GXM conjugates (which are
lyophilized), they preferentially bind to native wEPS. This trend may
reflect enhanced antigen presentation in the smaller native EPS parti­
cles. Furthermore, it is possible that similar effects occur when other
microbial polysaccharides are isolated by precipitation, lyophilization
and reconstitution techniques, which argues for caution in extrapolating
observations with different methods of preparation to those present in
native macromolecules. In a recent review, Yi and colleagues discussed
the dehydration of polysaccharides as one of the key procedures in
processing them and noted that vacuum drying and hot air drying both
lead to larger molecular weights, poorer solubility, and increased inci­
dence of aggregation compared to freeze drying (Yi et al., 2020). We

have observed each of these three effects upon lyophilization of
C. neoformans EPS. Similarly, increased apparent size by DLS, poor sol­
ubility, and increased aggregation have also been reported for poly­
saccharides from acorn (Ahmadi, Sheikh-Zeinoddin, Soleimanian-Zad,
Alihosseini, & Yadav, 2019), Chinese medicinal herb Bletilla striata
(Kong et al., 2015), mushroom Inonotus obliquus (Ma, Chen, Zhu, &
Wang, 2013), comfrey root (Shang et al., 2018), and finger citron fruits
(Wu, 2015). Further studies will be necessary to tease apart the effects of
isolation, freeze-, and vacuum-drying on polysaccharides, particularly
for cryptococcal EPS. Nevertheless, our observations together with
7


M.P. Wear et al.

Carbohydrate Polymers 291 (2022) 119547

Fig. 5. Structural model for the effects of lyophilization on C. neoformans EPS structure. EPS harvested from encapsulated C. neoformans is small, hydrated, rigid, and
ordered when purified in its native form. Lyophilization causes alterations to the native form, resulting in disordered, condensed conformations that exclude water.
Solvation of the lyophilized sample proceeds through gelation, wherein particles aquire a hydration shell but not structural waters. Over time in solution (28+ days)
the condensed conformation will be lost and a polymer-like structure more similar to, but not the same as, the native structure is adopted.

reports of other polysaccharides undergoing physicochemical alteration
upon dehydration (Yi et al., 2020) suggest that this may be a widespread
phenomenon for such polymers and argues for caution when interpret­
ing findings from rehydrated material.
In conclusion, scientists investigating the immunological properties
of cryptococcal polysaccharides should be aware that the method of
purification can affect its physicochemical properties, which in turn can
affect some of the immunological properties of polysaccharides. The

physicochemical alterations exacted by CTAB and lyophilization upon
polysaccharides could explain much of the variability in published
studies (Cordero et al., 2013; Crawford et al., 2020; Pierini & Doering,
2001; Zaragoza, Telzak, Bryan, Dadachova, & Casadevall, 2006) and
suggest the need for a renewed effort to characterize cryptococcal
polysaccharides using isolation techniques that maintain these mole­
cules in their native states.

(GOIPG/2016/998). The 600 MHz ssNMR facilities used in this work are
operated by The City College of New York and the City University of
New York Institute for Macromolecular Assemblies. The content is solely
the responsibility of the authors and does not necessarily represent the
official views of the National Institutes of Health.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.carbpol.2022.119547.
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Acknowledgments
The mechanism figure was created with Biorender software. We
thank Dr. Christine Chrissian for assistance with the ssNMR data
acquisition and analysis. We thank Barbara Smith and the Electron
Microscopy core facility at Johns Hopkins School of Medicine for their
analysis of the EPS samples as well as conversations about these data.
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